Investigating the ameliorative potential of Aspalathus linearis and Cyclopia intermedia against lipid accumulation, lipolysis, oxidative stress and inflammation

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Mokadi Peggy Mamushi

Thesis presented in fulfilment of the requirements for the degree of Masters in Science (Medical Physiology) in the Faculty of Medicine and Health Science at Stellenbosch University

SUPERVISOR: Dr C Pheiffer

CO-SUPERVISORS: Dr BU Jack and Prof SS Du Plessis

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i DECLARATION

By submitting this thesis electronically, I declare that the entirety of the work contained therein is my own, original work, that I am the authorship owner thereof (unless to the extent explicitly otherwise stated) and that I have not previously in its entirety or in part submitted it for obtaining any qualification.

……….. Miss Mokadi Peggy Mamushi

March 2020 ……… Date

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ii Despite the availability of several treatment regimens, obesity continues to be one of the greatest health challenges of the 21st century. In recent years, plant polyphenols have attracted increasing attention as nutraceuticals that are able to prevent or treat obesity and its co-morbidities. However, the first-line screening of these compounds is hampered by the shortage of in vitro experimental models that mimic the complex pathophysiology of obesity (excess lipid accumulation, basal lipolysis, inflammation and oxidative stress) in vivo. The aim of this study was two-fold. Firstly, establish a 3T3-L1 adipocyte in vitro model that more closely mimics obesity in vivo, and secondly to investigate the ameliorative properties of Aspalathus linearis, Cyclopia intermedia and their major polyphenols against these conditions.

Methods

For the experimental model, 3T3-L1 pre-adipocytes were differentiated in 5.5 mM, 25 mM or 33 mM glucose concentrations for 7 or 14 days. Lipid accumulation, basal lipolysis, oxidative stress, inflammation, mitochondrial activity and gene expression were assessed using Oil Red O staining, glycerol release, 2',7'-dichlorfluorescein-diacetate fluorescence to quantify reactive oxygen species, monocyte chemoattractant protein-1 secretion, the 3- [4, 5-Dimethylthiazol-2-yl]-2, 5 diphenyltetrazolium bromide assay and quantitative real time polymerase chain reaction, respectively. The ameliorative effects of Aspalathus linearis (Afriplex GRTTM) and Cyclopia

intermedia (CPEF) against these conditions were investigated by acute and chronic treatment of

the optimised experimental model with various concentrations of these plant extracts and their major polyphenol Aspalathin and Mangiferin respectively.

Results

Collectively lipid accumulation, basal lipolysis, oxidative stress, inflammation and expression of associated genes were higher after differentiation in 33 mM for 14 days compared to lower glucose concentrations and 7 days, thus these conditions were selected as the experimental model. Neither acute nor chronic treatment with 0.1 to 100 µg/ml of Aspalathus linearis and Cyclopia intermedia, and 0.1 to 100 µM of Aspalathin and Mangiferin significantly decreased lipid content. However, all treatments decreased basal lipolysis and increased mitochondrial activity.

Conclusion Background

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iii Differentiation of 3T3-L1 pre-adipocytes in 33 mM glucose for 14 days increased basal lipolysis, oxidative stress and inflammation compared to lower glucose concentrations and differentiation for 7 days. Aspalathus linearis, Cyclopia intermedia, Aspalathin and Mangiferin ameliorated the increased basal lipolysis under these conditions. This study showed that differentiation in 33 mM glucose for 14 days may offer potential as an experimental model that more closely mimics obesity

in vivo and may thus improve first-line screening for anti-obesity therapeutics. Aspalathus linearis, Cyclopia intermedia and their major polyphenol Aspalathin and Mangiferin, respectively may

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OPSOMMING

Agtergrond

Ten spyte van die beskikbaarheid van verskeie behandelingsregimes, bly vetsug steeds een van die grootste gesondheidsuitdagings van die 21ste eeu. In die onlangse verlede het polifenole, afkomstig vanaf plante, toenemende aandag getrek as funksionele voedingsmiddels wat vetsug en die ko-morbiditeit daarvan kan voorkom of behandel. Die eerste-lyn-sifting van hierdie verbindings word egter belemmer deur die tekort aan in vitro eksperimentele modelle wat die komplekse patofisiologie van vetsug (oortollige lipiedakkumulasie, lipolise, inflammasie en oksidatiewe stres) in vivo kan naboots. Die doel van hierdie studie was tweeledig. Die eerste doelwit was om 'n 3T3-L1-adiposiet in vitro-model op te stel wat vetsug in vivo naboots, en tweedens om die verligtingseienskappe van Aspalathus linearis, Cyclopia intermedia en hul belangrikste polifenole teen hierdie toestande te ondersoek.

Metodes

Vir die eksperimentele model is 3T3-L1 pre-adiposiete vir 7 of 14 dae in 5.5 mM, 25 mM of 33 mM glukosekonsentrasies gedifferensieer. Lipiedakkumulasie, lipolise, oksidatiewe stres, inflammasie, mitochondriale aktiwiteit en geenuitdrukking is onderskeidelik bepaal met behulp van “Oil Red O”-kleuring, gliserolvrystelling, 2', 7'-dichlorfluoresceïne-diasetaat fluoressensie om reaktiewe suurstofspesies te bepaal, monosiet chemo-aantrekkingskrag proteïen-1 sekresie, die 3- [4, 5-dimetieltiazol-2-yl]-2, 5 difenieltetrazoliumbromied-toets en kwantitatiewe reële tyd polimerase kettingreaksie. Die verbeteringseffekte van Aspalathus linearis (Afriplex GRTTM_{) en}

Cyclopia intermedia (CPEF) op hierdie toestande is ondersoek deur die akute en chroniese

behandeling van die geoptimaliseerde eksperimentele model met verskillende konsentrasies van hierdie plantekstrakte en hul belangrikste polifenole, Aspalathin en Mangiferin, onderskeidlik. Resultate

Lipiedakkumulasie, lipolise, oksidatiewe stres, inflammasie en uitdrukking van gepaardgaande gene was gesamentlik hoër na differensiasie in 33 mM glukose vir 14 dae in vergelyking met laer glukosekonsentrasies en 7 dae; dus is hierdie toestande as die eksperimentele model gekies. Nie akute of chroniese behandeling met 0,1 tot 100 µg/ml Aspalathus linearis en Cyclopia intermedia,

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v en 0,1 tot 100 µM Aspalathin en Mangiferin het die lipiedinhoud beduidend verlaag nie. Al die behandelings het egter lipolise verminder en mitochondriale aktiwiteit verhoog.

Gevolgtrekking

Differensiasie van 3T3-L1 pre-adiposiete in 33 mM glukose vir 14 dae het lipolise, oksidatiewe stres en inflammasie verhoog, vergeleke met laer glukosekonsentrasies en differensiasie vir 7 dae.

Aspalathus linearis, Cyclopia intermedia, Aspalathin en Mangiferin het die verhoogde lipolise

onder hierdie toestande verlig. Hierdie studie bewys dat die differensiasie in 33 mM glukose vir 14 dae potensiaal bied as 'n eksperimentele model wat vetsug in vivo naboots en sodoende eerste-lyn-sifting vir terapie teen vetsug kan verbeter. Aspalathus linearis en Cyclopia intermedia en hul belangrikste polifenole, Aspalathin en Mangiferin, toon dus potensiaal as antilipolitiese middels wat vetsug-geassosieerde lipolise kan verlig.

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ACKOWLEDGEMENTS

Firstly, I would like to thank our almighty God for if it was not by His grace and mercy, I would have not made it this far. He provided me with strength and protection during the research. Most importantly he blessed me with the humblest, loving, caring and beautiful souls that made my journey easy and educational.

I would also like to thank my main supervisor, Dr Carmen Pheiffer, for giving me the opportunity to do a NRF internship with her. I never thought that a girl from the deep rural area of Limpopo, who could not even speak one sentence in English could ever land an opportunity to work alongside the best researchers at the Biomedical Research and Innovation Platform of the South African Medical Research Council. Dr Pheiffer took me in with my flaws and made me into the scientist I am today. She was always patient and understanding even when she should not have been. She was a good mentor to me, ignited my love for research and sparked my interest to pursue a MSc under her supervision.

Thank you to my co-supervisor, Dr Babalwa Jack for her dedication towards my studies. She exceeded expectations, helping me with my experiments, presentations, funding applications and many more. Dr Jack trained me in Tissue Culture and was always prepared to work after hours and on weekends when needed. She taught me with love and compassion, and always remained calm and patient with me. I published my first review with her, with the little knowledge of laboratory work I had when I started my studies. I am greatly indebted to Dr Jack, she is an inspiration to me and someone that I will strive to become.

I would like to acknowledge Asive Myataza, whom I call my living angel. She was always there for me, even before I came to Cape Town. She hosted me upon my arrival in Cape Town and has been with me ever since. She helped me financially, physically, spiritually and emotionally. She became the sister I never had, she compliments me even when I know I failed big time and she boosted my confidence so much.

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vii Thank you to, Yoonus Ebrahim, Stephanie Dias and Tarryn Willmer, my team members (Epigenetics group), who also contributed to my development by providing stimulating discussions during our journal clubs and research meetings, and for helping me prepare for seminar presentations. I would especially like to thank Yoonus for training me in quantitative real time PCR.

I would also like to thank Prof Johan Louw for assisting with tuition fees.

I acknowledge Prof Stefan du Plessis, my co-supervisor, for his scientific input and support throughout this study.

Lastly, I would like to acknowledge the South African Medical Research Council (Research and Capacity Division), Stellenbosch University, Ethel and Ernst Eriksen trust, Harry Crossley and the National Research Foundation for funding and support (NRF-Grant- holder linked bursary- Grant number: 113459).

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4.2.1 Acute treatment ... 58 4.2.2 Chronic treatment... 58 4.3 Summary of results ...65 5. Discussion...68 5.1 Model development ...68 5.1.1 Lipid accumulation ... 68 5.1.2 Basal lipolysis ... 69 5.1.3 Oxidative stress ... 70 5.1.4 Inflammation ... 71 5.1.5 Mitochondrial activity ... 72 5.2 Treatment ...73 5.2.1 Lipid accumulation ... 73 5.2.2 Basal lipolysis ... 74 5.2.3 Mitochondrial activity ... 75

5.3 Strengths and limitations...75

5.4 Conclusion ...76

5.5 Future work ...77

6. Bibliography ...78

7. Appendix ...95

7.1 Aseptic technique ...95

7.2 Reagents and kits ...96

7.3 List of equipment and software ...98

7.4 Preparation of medium and buffers ...100

7.5 Assays ...102

7.5.1 Preparation of the ORO and CV stains ... 102

7.5.2 Preparation of the MTT ... 102

7.6 Treatments...103

7.7 Supplementary data ...104

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LIST OF TABLES

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Table 2.1 Classification of obesity and metabolic risk ... 8

Table 2.2 An overview of the current obesity therapeutic strategies ... 14

Table 3.1 Cell densities used for seeding 3T3-L1 pre-adipocytes ... 30

Table 3.2 Reaction components used for reverse transcription ... 40

Table 3.3 Reaction components for qRT-PCR reactions ... 42

Table 3.4 Taqman probes ... 42

Table 4.1 RNA concentrations, total yield and purity ... 53

Table 4.2 RNA integrity ... 54

Table 4.3 Assessment of genomic DNA contamination ... 55

Table 4.4 Amplification efficiency ... 55

Table 4.5 Results for model development ... 65

Table 4.6 Results of treatment ... 66

Table 7.1 List of reagents... 96

Table 7.2 List of kits ... 97

Table 7.3 List of equipment and consumables... 98

Table 7.4 List of software ... 99

Table 7.5 Preparation of medium... 100

Table 7.6 Preparation of DMEM without phenol red ... 100

Table 7.7 Sorenson's buffer ... 101

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LIST OF FIGURES

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Figure 2.1 Risk factors for obesity ... 10

Figure 2.2 Mechanisms relating adipocyte hypertrophy to metabolic disease ... 13

Figure 2.3 Chemical structure of Aspalathin ... 18

Figure 2.4 Chemical composition of Aspalathus linearis (Afriplex GRT) ... 19

Figure 2.5 Chemical structure of Mangiferin ... 20

Figure 2.6 Chemical composition of Cyclopia intermedia (CPEF) ... 21

Figure 3.1 Experimental overview ... 26

Figure 3.2 Cell counting ... 29

Figure 3.3 Experimental protocol for model development ... 31

Figure 3.4 Treatment experimental outline ... 33

Figure 4.1 Effect of glucose and differentiation times on lipid accumulation ... 46

Figure 4.2 Effect of glucose and differentiation times on basal lipolysis ... 47

Figure 4.3 Effect of glucose and differentiation times on oxidative stress ... 49

Figure 4.4 Effect of glucose and differentiation times on MCP1 secretion ... 50

Figure 4.5 Effect of glucose and differentiation times on mitochondrial activity ... 52

Figure 4.6 Effect of glucose and differentiation times on adipogenesis, lipid metabolism and adipokine genes ... 56

Figure 4.7 Effect of glucose and differentiation times on gene expression of oxidative stress markers ... 57

Figure 4.8 Effect of acute treatment on lipid content ... 59

Figure 4.9 Effect of acute treatment on glycerol release ... 60

Figure 4.10 Effect of acute treatment on mitochondrial activity ... 61

Figure 4.11 Effect of chronic treatment on lipid accumulation ... 62

Figure 4.12 Effect of chronic treatment on glycerol release ... 63

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LIST OF ABBREVIATIONS

ACACA Acetyl-CoA carboxylase alpha

ADM Adipogenesis inducing media

ADSCs Adipocyte derived stem cells

ADIPOQ Adiponectin

AMM Adipogenesis maintenance media

AT Adipose tissue

ATCC American type culture collection

BAT Brown adipose tissue

B2M Beta-2-microglobulin

BMI Body mass index

BSA Bovine serum albumin

cDNA complimentary DNA

CO2 Carbon dioxide

CPEF Crude polyphenol enriched fraction

CV Crystal violet

CVD Cardiovascular disease

DCF 2',7'-dichlorfluorescein

DCFH-DA 2',7'-dichlorfluorescein-diacetate

DEX Dexamethasone

DIO Diet induced obesity

DMSO Dimethyl sulfoxide

DNA Deoxyribonucleic acid

DPBS Dulbecco’s phosphate buffered saline

FA Fatty acid

FBS Foetal bovine serum

FDA Food and drug administration

GATA GATA-binding factor 2

GRT Green rooibos tea

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xiv HPLC High performance liquid chromatography

HSL Hormone sensitive lipase

HRP Horseradish peroxidase

Hrs Hours

IBMX 3-isobutyl-1-methylxanthine

IL6 Interleuken-6

Kg kilogram

LMIC Low- and middle-income countries

M Meters

MCP1 Monocyte chemotactic protein-1

Min Minutes

MSCs Mesenchymal stem cells

MRI Magnetic resonance imaging

MTT 3- [4, 5-Dimethylthiazol-2-yl]-2, 5 diphenyltetrazolium bromide NOX Nicotinamide adenine dinucleotide phosphate (NADPH) oxidase

NRF1 Nuclear respiratory factor 1

ORO Oil Red O

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PPARγ Peroxisome proliferator-activated receptor gamma qRT-PCR Quantitative real-time polymerase chain reaction

RNA Ribonucleic acid

ROS Reaction oxygen species

RPL13 Ribosomal protein L13a

RT Reverse transcription

SA South Africa

SAT Subcutaneous adipose tissue

Sec Seconds

spp. Species

SREBF1 Sterol regulatory element-binding transcription factor 1

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T75 75 cm2_{tissue culture flask}

TNFα Tumour necrosis factor alpha

UK United Kingdom

UCP1 Uncoupling protein-1

UN United Nations

USA United States of America

VAT Visceral adipose tissue

WAT White adipose tissue

WC Waist circumference

WHO World Health Organisation

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2

1. Introduction

1.1 Background

Obesity is a multifactorial disorder characterised by the excessive accumulation of fat to the extent that it negatively affects health (Hruby & Hu, 2015). Recent estimates show that approximately 641 million individuals worldwide (8.9%) are obese, with obesity rates projected to reach 20% by 2025 (NCD-RisC, 2016). Obesity increases the risk of developing chronic disorders such as insulin resistance, type 2 diabetes (T2D), cardiovascular disease (CVD) and cancer (Haslam & James, 2005). Although obesity was historically associated with developed countries, high rates are now reported in low- and middle-income countries (LMIC), driven by factors such as urbanisation, sedentary lifestyles and unhealthy diets (Ford, Patel & Narayan, 2017; Fox, Feng & Asal, 2019). Effective interventions are required to reduce the burden of obesity on health systems, particularly those in LMIC that are already over-burdened and under-resourced and least able to respond to the escalating obesity crisis.

The mechanisms that underly the development of obesity and its complications are not yet fully elucidated, although several studies show that excessive energy intake leads to adipose tissue enlargement by hypertrophy (Jo et al., 2009; Sun, Kusminski & Scherer, 2011; Jung & Choi, 2014). Adipocyte hypertrophy is associated with increased lipid accumulation, basal lipolysis, oxidative stress and inflammation. Excessive lipid accumulation and basal lipolysis leads to fatty acid (FA) secretion, with harmful effects on peripheral tissues such as the liver and muscle where it induces insulin resistance (Boden, 2011; Wang, Scherer & Gupta, 2014). Furthermore, adipocyte hypertrophy induces reactive oxygen species (ROS) and oxidative stress (Boden, 2011; Wang, Scherer & Gupta, 2014). In addition, adipocyte hypertrophy is associated with chronic low-grade inflammation (Shoelson, Herrero & Naaz, 2007; Boutens & Stienstra, 2016), which is mediated by the increased secretion of monocyte chemotactic protein (MCP1). This cytokine attracts pro-inflammatory M1 macrophages to adipose tissue and decreases secretion of anti-pro-inflammatory adipokines such as adiponectin (Jung & Choi, 2014). Together, all these co-morbidities of obesity induce metabolic disease (Jung & Choi, 2014; Gambero & Ribeiro, 2015).

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3 Despite the availability of several anti-obesity strategies such as lifestyle modifications (mainly diet and exercise), pharmaceutical compounds and surgery for the morbidly obese, there is still a lack of safe, effective and long-term therapies. In recent years, plant polyphenols have attracted increasing attention as nutraceuticals that are able to prevent or treat obesity and its co-morbidities (Sun, Wu & Chau, 2016). Aspalathus linearis and Cyclopia species, more commonly known as rooibos and honeybush respectively, are indigenous South African plants that are widely consumed as herbal teas due to their pleasant aroma and taste (Joubert et al., 2008). Furthermore, these herbal teas are attracting increased interest due to their health promoting properties. Previous studies have shown that they are able to prevent the development of obesity (Dudhia et al., 2013; Pheiffer et al., 2013; Sanderson et al., 2014; Jack et al., 2017), insulin resistance (Mazibuko et al., 2013), T2D (Muller et al., 2012; Chellan et al., 2014) and CVD (Dludla et al., 2014). Moreover, a dihydrochalcone C-glycoside, Aspalathin, which is the main flavonoid of rooibos was shown to improve lipid metabolism in insulin resistant 3T3-L1 adipocytes (Mazibuko et al., 2015), while Mangiferin, a xanthone C- glycoside from honeybush was shown to ameliorate insulin resistance by inhibiting inflammation and regulating adipokine secretion in adipocytes cultured under hypoxic conditions (Yang et al., 2017).

1.2 Problem statement

Obesity is considered one of the greatest health challenges of the 21st_{century. Obesity increases}

the risk of developing chronic metabolic diseases such T2D, CVD and cancer, thereby significantly decreasing life expectancy (Peeters et al., 2003; Haslam & James, 2005). Effective interventions are required to reduce the burden of obesity on health systems, particularly those in LMIC that are already over-burdened and under-resourced and least able to respond to the escalating obesity crisis. Lifestyle modifications such as improved diet and physical activity are the most effective anti-obesity strategies (Lagerros & Rössner, 2013). However, these lifestyle modifications are difficult to adhere to, thus increasing reliance on therapeutics, which unfortunately are plagued by several side effects (Sweeting, Hocking & Markovic, 2015). There is an urgent need to identify more effective and safer anti-obesity treatments.

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4 1.3 Rationale

In recent years, plant polyphenols have attracted increasing attention as nutraceuticals that are able to prevent or treat obesity and its co-morbidities. In particular, Aspalathus linearis, Cyclopia

intermedia and their major polyphenol Aspalathin and Mangiferin, respectively have been shown

to possess ameliorative properties against these conditions (Muller et al., 2012; Dudhia et al., 2013; Mazibuko et al., 2013; Pheiffer et al., 2013; Chellan et al., 2014; Dludla et al., 2014; Sanderson et al., 2014; Jack et al., 2017; Yang et al., 2017). The use of animal models to screen these compounds are restricted due to ethnical concerns, thus, in vitro experimental models are widely used as first-line screening tools (Nilsson et al., 2012; Denayer, Stöhrn & Van Roy, 2014; Barrett, Mercer & Morgan, 2016). However, in vitro models do not reflect the complex pathophysiology of obesity (excess lipid accumulation, basal lipolysis, inflammation and oxidative stress) in vivo, which may hamper bioactivity testing (Ruiz-Ojeda et al., 2016; Langhans, 2018). It is therefore imperative to develop an in vitro model that mimics the pathophysiology of obesity in vivo (excess lipid accumulation, basal lipolysis, inflammation and oxidative stress) in order to improve the screening of anti-obesity therapeutics.

1.4 Hypothesis

1. We hypothesised that differentiating 3T3-L1 adipocytes in higher glucose concentrations for an extended period of time (compared to standard culture conditions) will exacerbate the co-morbidities associated with obesity (increased lipid accumulation, basal lipolysis, oxidative stress and inflammation).

2. Treatment with Aspalathus linearis, Cyclopia intermedia and their major polyphenols (Aspalathin and Mangiferin) will have ameliorative effects against increased lipid accumulation, basal lipolysis, oxidative stress and inflammation.

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5 1.5 Aims

The aims of this study are two-fold.

• To develop a 3T3-L1 adipocyte in vitro model that more closely mimics obesity in vivo (increased lipid accumulation, basal lipolysis, oxidative stress and inflammation compared to standard conditions).

• To assess the ameliorative effects of Aspalathus linearis, Cyclopia intermedia and their major polyphenol Aspalathin and Mangiferin, respectively, against increased lipid accumulation, basal lipolysis, oxidative stress and inflammation.

1.6 Objectives Aim 1

• To differentiate 3T3-L1 pre-adipocytes in 5.5 mM, 25 mM or 33 mM glucose concentrations for 7 or 14 days; and

• To assess lipid accumulation, basal lipolysis, oxidative stress, inflammation, mitochondrial activity and gene expression.

Aim 2

• Acute and chronic treatment of the optimised model with Aspalathus linearis (Afriplex GRTTM), Cyclopia intermedia (CPEF) and their major compound Aspalathin and Mangiferin, respectively; and

• To assess lipid accumulation, basal lipolysis, oxidative stress, inflammation, mitochondrial activity and gene expression in the treated adipocytes.

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2. Literature review

2.1 Obesity definition

The World Health Organisation (WHO) defines obesity as a complex metabolic disease characterised by the excessive accumulation of body fat to the extent that it negatively affects health (WHO, 2018). The body mass index (BMI) is the most commonly used method for assessing obesity; it is calculated by dividing an individual's weight in kilograms (kg) by the square of their height in meters (m) (kg/m2) (Nuttall, 2015). The WHO guidelines for the classification of obesity and metabolic risk are shown in Table 2.1 (WHO, 2000). However, the use of BMI to assess obesity is widely criticised. BMI is affected by gender and ethnicity and is not able to discriminate between muscle and fat mass. Methods that have been recommended as an alternative to BMI include waist circumference (WC), waist to hip ratio (WHR) and skinfold thickness (Kuriyan, 2018; Osayande, Azekhumen & Obuzor, 2018; Eghan et al., 2019). Values of WC ≥ 88 cm for women and ≥ 102 cm for men are associated with a high metabolic risk (Table 2.1) (Kuriyan, 2018). WHR measures the ratio of the WC to the hip circumference; ratios ≥ 0.80 for women and ≥ 0.95 for men are associated with metabolic risk (Table 2.1). Skinfold thickness measures the thickness of subcutaneous tissue (skin layer) at sites such as the triceps, biceps or right hipbone with specialised callipers (Cornier et al., 2011; Kuriyan, 2018). More sensitive, but costly and technically challenging techniques due to their reliance on specialised equipment, include bioelectrical impedance analysis, computed tomography, dual-energy X-ray absorptiometry, magnetic resonance imaging and underwater weighing (Borga et al., 2018; Cornier et al., 2011; Kuriyan, 2018; Lee et al., 2018).

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8 Table 2.1 Classification of obesity and metabolic risk

Table taken from WHO, 2000.

2.2 Epidemiology of obesity 2.2.1 Global prevalence

Globally, the prevalence of obesity has nearly tripled since 1975 (WHO, 2018). Recent estimates show that approximately 641 million individuals worldwide (8.9%) are obese, with the prevalence of obesity projected to reach 20% by 2025 (NCD-RisC, 2016). According to the WHO, approximately 41 million children under the age of five years were overweight or obese in 2016. The prevalence of overweight and obesity dramatically increased from 4% in 1975 to 18% in 2016 among children and adolescents aged between five and nineteen years (WHO, 2018).

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9 2.2.2 Prevalence in South Africa

Although obesity was historically associated with developed countries, in recent years, high rates of obesity are reported in LMIC (Ng et al., 2014; Ford, Patel & Narayan, 2017), thus placing a major burden on the already struggling and over-burdened health systems of these countries. Between 1980 and 2015, the prevalence of overweight and obesity has nearly doubled in Africa (Chooi, Ding & Magkos, 2019). South Africa, a middle-income country at the southernmost tip of Africa, has the highest rates of obesity in Africa. In 2013, approximately 69.3% of women older than 20 years were overweight, of whom 42% were obese. Although rates were lower in men (38.8% overweight and 13.5% obese), they are nevertheless higher than the global average (Ng et al., 2014). Similar trends were also observed in children and adolescents, with obesity rates of 9.6% and 7.0% in girls and boys respectively.

2.3 Risk factors for obesity

Obesity is a multifactorial disease caused by a prolonged positive energy imbalance, resulting from increased energy intake and reduced energy expenditure (Hill, Wyatt & Peters, 2012). Obesity occurs due to the interplay of a number of factors including diet, physical activity, age, genetics and epigenetics (Figure 2.1), although the consumption of high calorie diets and the lack of physical activity are considered the main drivers of the current obesity pandemic (Hill, Wyatt & Peters, 2012). Furthermore, societal factors including socio-economic status and urbanisation have also contributed to the increased prevalence of obesity in developing countries (Micklesfield et al., 2013). Other risk factors for obesity include medications such as antidepressants, glucocorticoids, psychological factors and neuroendocrine-related factors (Grundy et al., 2014; Thaker, 2017; Blüher, 2019).

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Figure 2.1 Risk factors for obesity

Overweight and obesity results from an energy imbalance due to the interplay of a multitude of risk factors.

2.4 Adipose tissue

Adipose tissue is a loose connective tissue that is mainly composed of adipocytes or fat cells, although other cell types present include fibroblasts, mesenchymal stem cells (MSCs), macrophages and vascular endothelial cells (Esteve Ràfols, 2014; Lynes & Tseng, 2018; Luong, Huang & Lee, 2019). Adipose tissue is widely distributed throughout the body and represents 15-20% of body weight in normal-weight healthy men, and 25-30% in normal-weight healthy women (Gallagher et al., 2000). There are two types of adipose tissue, namely white adipose tissue (WAT) and brown adipose tissue (BAT), which vary according to location, cellular structure and physiological function (Saely, Geiger & Drexel, 2012; Lee, Mottillo & Granneman, 2014).

2.4.1 White adipose tissue

The WAT is the most common type of adipose tissue, composed of densely packed mature adipocytes (35-75%), pre-adipocytes, MSCs, T regulatory cells, endothelial precursor cells and macrophages (Esteve Ràfols, 2014; Lynes & Tseng, 2018; Luong, Huang & Lee, 2019). Adipocytes within WAT are spherical in morphology and contain a single large lipid droplet that occupies up to 85% of the cell, a flattened peripheral nucleus and few mitochondria in the

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11 periphery of the cytoplasmic area (Tandon, Wafer & Minchin, 2018). The main function of WAT is to regulate energy balance by storing excess energy as triacylglycerol and releasing energy in the form of FAs into the circulation when needed (Cinti, 2005; Church, Horowitz & Rodeheffer, 2012). WAT expands by two mechanisms, namely hypertrophy which increases adipocyte size, or hyperplasia, which increases adipocyte number (Jo et al., 2009). Adipocyte hypertrophy is implicated in the pathogenic mechanisms underlying obesity (Figure 2.2) (Lee, Wu & Fried, 2010; Sun, Kusminski & Scherer, 2011; Kahn, Wang & Lee, 2019; Longo et al., 2019). Adipocyte hypertrophy leads to increased basal lipolysis and FA secretion, increased inflammation through secretion of MCP1 and pro-inflammatory macrophage infiltration, and increases ROS and oxidative stress, conditions implicated in the pathophysiological mechanisms of metabolic disorders (Figure 2.2) (Boden, 2011; Sun, Kusminski & Scherer, 2011; Jung & Choi, 2014; Boutens & Stienstra, 2016). Obesity increases the risk of T2D, CVD and several types of cancer (Haslam & James, 2005). Globally, overweight and obesity contributes to about 44%, 23% and 7– 41% of T2D, CVD and cancers, respectively (Frühbeck et al., 2013). Paradoxically, not all obese individuals develop chronic disease. Approximately 10-25% of obese individuals are metabolically healthy, while a similar percentage of normal weight individuals are metabolically unhealthy and have an increased risk of developing chronic disease (Blüher, 2010; Denis & Obin, 2013; Smith, Mittendorfer & Klein, 2019). This suggests that fat distribution rather than fat mass define metabolic risk (Goossens, 2017; Kwon, Kim & Kim, 2017; Grundy, Williams & Vega, 2018), and underscores the role of genetics and environmental factors in the development of obesity and metabolic disorders.

Other than being a fat reservoir, WAT also functions as a cushion to protect vital organs against mechanical stress and acts as an insulator, which controls heat conduction through the skin (Zwick et al., 2018). Emerging evidence suggest that WAT is a major endocrine organ that plays a role in regulating whole-body metabolic homeostasis via the secretion of adipokines (Kershaw & Flier, 2004; Coelho, Oliveira & Fernandes, 2013). These molecules play a vital role in regulating physiological processes such as appetite, energy balance, lipid and glucose metabolism and systemic immunity (Coelho, Oliveira & Fernandes, 2013; Musi & Guardado-Mendoza, 2014; Booth et al., 2016; Choe et al., 2016). WAT is mainly distributed within visceral adipose tissue

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12 (VAT) and subcutaneous adipose tissue (SAT) depots (Choe et al., 2016; Mittal, 2019). VAT is characterised by ectopic fat deposition around organs and is associated with higher metabolic risk. Several lines of evidence show that increased accumulation of VAT leads to impaired insulin action, increased inflammation and oxidative stress, and the development of metabolic disease (Alexopoulos, Katritsis & Raggi, 2014; Janochova, Haluzik & Buzga, 2019; Longo et al., 2019). Conversely, SAT is characterised by fat distribution in the femoral, hips and gluteal regions and is considered metabolically benign. SAT is a metabolic buffer that prevents lipotoxicity induced lipid overflow and ectopic fat accumulation in other tissues (Ibrahim, 2010). Inadequate SAT expansion leads to visceral and ectopic fat deposition, adipokine dysregulation and insulin resistance (Longo et al., 2019).

2.4.2 Brown adipose tissue

In contrast to WAT that functions as an energy reservoir, BAT induces non-shivering thermogenesis (Keipert & Jastroch, 2014; Jastroch, Oelkrug & Keipert, 2018). Adipocytes within BAT are smaller and contain many mitochondria with increased expression of mitochondrial uncoupling protein 1 (UCP1), which gives BAT its unique ability to generate heat via adaptive thermogenesis (Saely, Geiger & Drexel, 2012; Keipert & Jastroch, 2014; Lee, Mottillo & Granneman, 2014; Jastroch, Oelkrug & Keipert, 2018). BAT is mainly present in babies and decreases with age (Yoneshiro et al., 2011; Gonçalves et al., 2017; Zoico et al., 2019). Beige adipocytes are a relatively newly discovered adipocyte found interspersed within white adipocytes (Wu et al., 2012; Sepa-Kishi & Ceddia, 2018; Lizcano, 2019; Zoico et al., 2019). Under basal conditions, beige adipocytes are morphologically indistinguishable from white adipocytes, although they share similar functions with both white and brown adipocytes (Sepa-Kishi & Ceddia, 2018). Beige adipocytes are activated upon cold exposure, chronic endurance exercise or β3-adrenergic stimulation, and subsequently assume a brown adipocyte-like morphology with higher mitochondria content and multiple small lipid droplets (Sepa-Kishi & Ceddia, 2018). Activated beige adipocytes have higher expression of UCP1 and enhanced thermogenic capacity, thus leads to improved lipid and glucose metabolism (Wu et al., 2012; Sepa-Kishi & Ceddia, 2018; Lizcano, 2019).

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13 Figure 2.2 Mechanisms relating adipocyte hypertrophy to metabolic disease

Healthy adipose tissue is characterised by anti-inflammatory M2 macrophages. Factors such as unhealthy diets, physical inactivity, genetics and stress leads to adipocyte hypertrophy. Hypertrophied adipocytes recruit pro-inflammatory M1 macrophages, increasing inflammation and oxidative stress and impairing lipid metabolism. These conditions have systemic effects on peripheral tissues such as the heart, liver, muscle, pancreas and brain, leading to the development of chronic diseases. Arrow up (↑) indicates increase and arrow down (↓) signifies decrease. Figure taken and modified from (Jack et al., 2019).

2.5 Obesity intervention

Current intervention strategies for obesity include lifestyle modification, pharmacotherapy and surgery for the morbidly obese (Table 2.2).

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14 Table 2.2 An overview of the current obesity therapeutic strategies

Treatment Strategy

Therapy

Mechanism/Activity References

Lifestyle modification Diet

Prevents excessive fat accumulation

(Fock & Khoo, 2013; Lagerros & Rössner, 2013)

Physical activity Increase metabolic rate

Pharmacological drugs _Orlistata

(Xenical, Alli)

Reduces fat absorption (pancreatic lipase inhibition) (Adan, 2013; Sweeting, Hocking & Markovic, 2015; Wharton, 2016; Patel & Stanford, 2018)

Lorcaserina (Belviq)

Appetite suppressant and promotes satiety (5-HT2c receptor agonist)

Phentermine/topiramate ERa (Qsymia)

Appetite suppressant (noradrenalin releaser and anticonvulsant/ neurostabilizer) Phentermine HClb (Adipex, Lomaira) Naltrexone SR/buproprion SRa (Contrave) Liraglutidea (Saxenda)

Appetite suppressant (noradrenalin releaser/sympaticomimetic) Appetite suppressant

(norepinephrine/dopamine reuptake inhibitor and opioid receptor antagonist) Appetite suppressant (GLP-1 receptor agonist)

Weight-loss surgery/ Bariatric surgeryc

Gastric sleeve; Adjustable gastric band; Roux-en-Y gastric bypass; Biliopancreatic diversion with

duodenal switch

Restricts food intake and reduces nutrient absorption by decreasing stomach size or changing the anatomy of gastrointestinal area; Appetite suppression due to physiological or hormonal changes; Increases energy expenditure (Kissler & Settmacher, 2013; Piché et al., 2015; Kassir et al., 2016; Wolfe, Kvach & Eckel, 2016)

a_{Approved by FDA for long-term use.}

b_{Approved by FDA only for short term use (≤ 12 weeks) and low dose.} c_{Only recommended for morbidly obese (BMI ≥ 40 kg/m}2_).

Table abbreviations: β3-AR, beta-3 adrenergic receptor; ER, extended release; FDA, Food and Drug Administration; GLP-1, glucagon-like peptide 1; 5-HT2c, serotonin receptor; SR, sustained release; HCl, hydrochloride. Table taken and modified from (Jack et al., 2019).

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15 2.5.1 Lifestyle modification

Lifestyle modifications such as calorie restriction and increased physical activity are the most effective first-line treatment strategies used in the management of overweight and obesity and have both been shown to induce significant weight loss (Table 2.2) (Fock & Khoo, 2013; Lagerros & Rössner, 2013). A low-calorie diet (800-1500 kcal/day) was shown to decrease bodyweight by 10% in obese individuals over a period of 3-12 months, while a very low-calorie diet (≤ 800 kcal/day) induced a 21.3% weight loss over a period of 6 months (Anderson, Luan & Høie, 2004; Fock & Khoo, 2013). A combination of diet and physical activity induced a 10.4% weight loss compared to diet (9.1%) or physical activity (2.1%) alone (Wing et al., 1998). However, studies have shown variability in an individual’s response to diet and exercise, which is most likely due to genetic variation (Bray, 2008). Furthermore, despite the success of lifestyle modification, adherence is poor, increasing reliance on pharmacological agents to manage obesity.

2.5.2 Pharmacotherapy

Pharmacotherapy is prescribed to individuals who are unable to control obesity with lifestyle modifications (Adan, 2013; Sweeting, Hocking & Markovic, 2015; Wharton, 2016; Patel & Stanford, 2018). Orlistat was approved by the Food and Drug Administration (FDA) for the long-term treatment and management of obesity in 1990 and is the oldest anti-obesity drug on the market (Hvizdos & Markham, 1999; Sweeting, Hocking & Markovic, 2015; Patel & Stanford, 2018). It inhibits pancreatic lipase, thus decreasing intestinal fat absorption and promoting faecal fat excretion (Hvizdos & Markham, 1999). However, Orlistat has several gastrointestinal adverse effects such oily stools, flatulence, increased defaecation, while some studies have shown that Orlistat interferes with nutrient and drug absorption (Adan, 2013; Patel & Stanford, 2018). Other anti-obesity drugs, including Lorcaserin, Phentermine/topiramate, Naltrexone/buproprion and Liraglutide have been approved by the FDA over the past few years and are currently used for the long-term treatment and management of obesity, while Phentermine hydrochloride is approved by the FDA only for short term use and at low dose (Table 2.2) (Adan, 2013; Sweeting, Hocking & Markovic, 2015; Wharton, 2016; Patel & Stanford, 2018). Several new drugs or drug combination therapies that target the central nervous system, the gastrointestinal tract or adipose tissue metabolism (either preventing the pathophysiology associated with adipose tissue dysfunction or

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16 promoting energy expenditure via thermogenesis) are currently under preclinical investigation or in clinical development and may be approved for obesity treatment in the next few years (Barja-Fenández et al., 2014; Kakkar & Dahiya, 2015).

2.5.3 Bariatric surgery

Bariatric surgery, a collective term for gastrointestinal surgical procedures performed on severely obese patients who have at least one obesity-associated co-morbidity (Table 2.2), is considered the most effective treatment for reducing excess body weight (Kissler & Settmacher, 2013; Piché et al., 2015; Wolfe, Kvach & Eckel, 2016). The currently established bariatric procedures include the sleeve gastrectomy, adjustable gastric band, laparoscopic Roux-en-Y gastric bypass, and biliopancreatic diversion with duodenal switch (Table 2.2) (Kissler & Settmacher, 2013; Piché et al., 2015; Wolfe, Kvach & Eckel, 2016). These surgical procedures restrict food intake and reduce nutrient absorption by decreasing stomach size or changing the anatomy of the gastrointestinal area, thus suppressing appetite and improving weight loss and metabolic status (Table 2.2) (Kissler & Settmacher, 2013; Piché et al., 2015; Wolfe, Kvach & Eckel, 2016). Despite the beneficial effects of bariatric surgery, these surgical procedures are expensive and are associated with post-surgery complications such as the malabsorption of essential vitamins, minerals and pharmacological drugs (Kassir et al., 2016).

2.6 Natural products as anti-obesity agents

Plant-derived natural products are attracting increasing interest as therapeutic agents due to the perception that they are safer and more cost-effective than synthetic drugs (Mopuri & Islam, 2017; Jack et al., 2019). Natural products from different sources, especially from plants, have been used as remedies for human diseases for centuries and offer potential as a source for the development of new drugs (Veeresham, 2012; Mushtaq et al., 2018). Polyphenols are secondary plant metabolites with a variety of beneficial health effects (Pandey & Rizvi, 2009). They are found in a variety of dietary sources including fruits, vegetables and beverages such as tea or wine and are widely explored as an alternative or as an adjunct to conventional obesity treatment (Manach et al., 2004; Pandey & Rizvi, 2009; Meydani & Hasan, 2010; Wang et al., 2014). Aspalathus linearis

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17 (rooibos) and Cyclopia ssp. (honeybush) are endemic South African plants consumed as herbal teas globally. These teas are caffeine-free and have a low tannin content (Joubert et al., 2008; Stander, Joubert & De Beer, 2019). Over the last few years, the global demand for these teas has increased and is partly attributed to their perceived beneficial health properties (Muller et al., 2018; Joubert et al., 2019). Likewise, studies have reported that the major phenolic compounds present in rooibos and honeybush possess anti-obesity effects (Fomenko & Chi, 2016; Johnson et al., 2018; Jack et al., 2019).

2.6.1 Aspalathus linearis

Aspalathus linearis (Brum.f) Dahlg. (Fabaceae), commonly known as rooibos, is a shrub-like

leguminous member of the fynbos biome indigenous to the Western and Northern Cape region of South Africa (Joubert et al., 2008; Joubert & de Beer, 2011). Rooibos is consumed as a “fermented” (oxidised) or “unfermented” (green) rooibos tea (Joubert & de Beer, 2011). During fermentation, the leaves and stems of the rooibos plant material are bruised and soaked to enhance the natural oxidation process, which gives rise to a rooibos infusion with a distinctive reddish-brown colour and a pleasant, slightly sweet flavour. Unfermented rooibos tea maintains its green colour due to limited oxidation (Joubert et al., 2008; Joubert & de Beer, 2011).

Unlike the fermented rooibos, which has reduced anti-oxidant content as a result of oxidation, the unfermented rooibos product preserves the quality of anti-oxidants and polyphenols such as Aspalathin (Marnewick et al., 2005; Villaño et al., 2010; Joubert & de Beer, 2011). Unfermented rooibos tea displays a 28% higher in vitro anti-oxidant capacity than fermented rooibos tea (Villaño et al., 2010). Aspalathin, a C-glucosyl dihydrochalcone unique to rooibos (Figure 2.3), is considered the major biological active polyphenol in rooibos (Joubert & de Beer, 2011). Other major bioactive compounds found in rooibos include the dihydrochalcone, Nothofagin, flavones (Orientin, Isoorientin, Luteolin, and Apigenin) and the flavonols (Quercetin and Rutin) (Joubert & de Beer, 2011). Phenylpyruvic acid-2-O-β-D-glucoside (PPAG) is one of the major constituents of fermented rooibos infusions (Muller et al., 2013).

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18 Figure 2.3 Chemical structure of Aspalathin

Chemical structure obtained from ChemSpider database (http://www.chemspider.com, 2019a).

In addition to its widely reported anti-oxidant properties (Joubert et al., 2005; Villaño et al., 2010), many studies have reported that rooibos tea has anti-mutagenic (Marnewick, Gelderblom & Joubert, 2000; Standley et al., 2001), anti-cancer (Marnewick et al., 2005), anti-inflammatory (Baba et al., 2009), cardioprotective (Pantsi et al., 2011; Dludla et al., 2014), anti-diabetic (Muller et al., 2012; Mazibuko et al., 2013; Kamakura et al., 2015; Sasaki, Nishida & Shimada, 2018), and anti-obesity (Beltrán-Debón et al., 2011; Sanderson et al., 2014) properties. Sanderson et al. showed that rooibos inhibits adipogenesis and intracellular lipid accumulation in 3T3-L1 adipocytes, accompanied by decreased expression of adipogenesis and lipid accumulation genes and proteins (Sanderson et al., 2014). Although the anti-obesity effects of Aspalathin have not been reported yet, this compound was shown to improve glucose and lipid metabolism in insulin resistant 3T3-L1 adipocytes (Mazibuko et al., 2015). Recently, a green rooibos extract (GRT) was manufactured by Afriplex (Paarl, Western Cape, SA) according to standardised conditions. The polyphenol content is enriched as the rooibos is not fermented (Figure 2.4). GRT was shown to have beneficial health effects against hyperglycaemia, oxidative stress and dyslipidaemia in high-fat diet-induced diabetic vervet monkeys (Orlando et al., 2019).

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19 Figure 2.4 Chemical composition of Aspalathus linearis (Afriplex GRT)

Figure shows high performance liquid chromatography (HPLC) chromatograms of the GRT extract (A) with compound content values presented in g/100g extract (B). Adapted and modified from (Patel et al., 2016).

2.6.2 Cyclopia spp.

Cyclopia spp. (Genus: Cyclopia Vent.; Family: Fabaceae; Tribe: Podalrieae) are indigenous South

African plants that are used to produce honeybush tea (Joubert et al., 2011, 2019). Twenty-three species of Cyclopia have been described thus far, with six species (C. subternata, C. genistoides,

C. intermedia, C. maculata, C. longifolia and C. sessiliflora) used for commercial tea production

(Joubert et al., 2011). Over the past 20 years, the honeybush tea industry has gained increased popularity, mainly due to its distinctively sweet flavour and aroma (Joubert et al., 2019). Plant material is fermented or oxidised at a high temperature to give the distinctive characteristics of honeybush tea. In recent years, unfermented or green honeybush has attracted interest due to its higher polyphenol content and perceived increased health effects (Joubert et al., 2011).

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20

Cyclopia spp. are a rich source of complex bioactive phenolic compounds, and high quantities of

the xanthones, Mangiferin (Figure 2.5) and its isomer, Isomangiferin, the flavanone, Hesperidin, the benzophenone glucosides (3-β-D-glucopyranosyliriflophenone, 3-β-D-glucopyranosyl-4-β-D -glucopyranosyloxyiriflophenone and 3-β-D-glucopyranosylmaclurin), as well as dihydrochalcone glycosides (Phloretin-3′,5′-di-C-β-D-glucoside and 3-hydroxy-phloretin-3′,5′-di-C-hexoside) have been detected in extracts of several Cyclopia plants (Jack et al., 2019; Joubert et al., 2019). Other phenolic compounds found in Cyclopia spp. include the flavanones (Hesperitin, Naringenin, Eriocitrin, Neoponcirin and Eriodictyol) and flavones (Luteolin, Scolymoside and Vicenin-2), which have been detected in relatively small amounts in extracts of several Cyclopia plants (Jack et al., 2019; Joubert et al., 2019).

Figure 2.5 Chemical structure of Mangiferin

Chemical structure obtained from ChemSpider database (http://www.chemspider.com, 2019b).

Cyclopia spp. exhibit potential anti-obesity (Dudhia et al., 2013; Pheiffer et al., 2013; Jack et al.,

2017, 2018), anti-diabetic (Muller et al., 2011; Chellan et al., 2014), anti-oxidant (Joubert et al., 2008; Lawal, Davids & Marnewick, 2019), mutagenic (van der Merwe et al., 2006), anti-cancer (Marnewick et al., 2005), phytoestrogenic (Mortimer et al., 2015), anti-osteoclastogenic (Visagie et al., 2015), anti-allergic (Murakami et al., 2018) and anti-inflammatory (Lawal, Davids & Marnewick, 2019) activities. Recently, Jack et al. demonstrated that a crude

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polyphenol-21 enriched organic fraction of C. intermedia (CPEF) possesses anti-obesity effects in vitro and in

vivo (Jack et al., 2017). The polyphenolic profile and compounds within CPEF are shown in Figure

2.6. Several studies have reported that Mangiferin, the major polyphenol detected within Cyclopia spp. (Figures 2.5 and 2.6), possesses anti-obesity properties in vitro in adipocyte models (Yoshikawa et al., 2002; Subash-Babu & Alshatwi, 2015; Yang et al., 2017). In vivo studies revealed that Mangiferin can reduce body weight and ameliorate obesity-associated metabolic conditions (Guo et al., 2011; Niu et al., 2012; Apontes et al., 2014; Lim et al., 2014; Acevedo et al., 2017; Wang et al., 2017).

Figure 2.6 Chemical composition of Cyclopia intermedia (CPEF)

Figure shows a HPLC chromatogram of CPEF (A) with compound content values presented in g/100g fraction (B). Adapted and modified from (Jack et al., 2017).

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22 2.7 Experimental models

Development of anti-obesity therapeutics require extensive preclinical screening which makes use of in vivo animal models and in vitro cell culture models.

2.7.1 In vivo models

Animal models have played a key role in elucidating the aetiology of obesity, including the physiological processes and genetic mechanisms that regulate energy homeostasis, appetite and adipose tissue metabolism (Agahi & Murphy, 2014; Barrett, Mercer & Morgan, 2016). In vivo models such as rodents and non-human primates have been used to better elucidate the efficacy, safety, pharmacology and pharmacokinetic potential of anti-obesity drug candidates before testing in humans (Agahi & Murphy, 2014). The currently used animal models of obesity include diet-induced obese (DIO) animal models and genetic models (Barrett, Mercer & Morgan, 2016). The DIO animal model involves feeding animals a diet that is high in fat, sucrose or fructose (or the combination) to induce obesity and several glucose and lipid metabolic abnormalities. As such, DIO models have been used to develop anti-obesity drugs because they provide a better understanding of the pathophysiology of obesity, especially the gene–environment interactions underlying the major causes of human obesity (Barrett, Mercer & Morgan, 2016; Ohta, Murai & Yamada, 2017). Genetic models have a spontaneous mutation or are genetically engineered to express certain physiological traits such as extreme obesity. They have been used to provide valuable insight into the pathophysiological contribution of particular genes to obesity (Agahi & Murphy, 2014; Barrett, Mercer & Morgan, 2016). The use of animal models for screening anti-obesity drugs is restricted due to ethical concerns and legislations including the “Three Rs” (reduction, refinement, and replacement) (Sharma et al., 2011; Pasupuleti, Molahally & Salwaji, 2016). As a result, in vitro models are commonly used as an alternative first line screening tool.

2.7.2 In vitro models

In vitro models such as immortalized cell lines are used as an alternative to in vivo animal testing

in biomedical research (Chacon et al., 1996, Romero et al., 2014). These in vitro models are

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23 understand their effects on cellular physiology and metabolism, dose response and toxicity effects (Astashkina, Mann & Grainger, 2012; Ruiz-Ojeda et al., 2016). In addition, in vitro models are useful for rapid and inexpensive screening of bioactive compounds in a controlled environment (temperature, pH, hormone and nutrient concentrations, etc.), thus reducing experimental variation. Cell lines provide homogeneous, genetically identical cellular populations and cell types which are at the same differentiation stage, therefore, allowing a homogeneous response to treatments and providing consistent and reproducible results (Carter & Shieh 2010). However, cell lines have altered characteristics and functions compared to their in vivo counterparts.

In vitro models such as 3T3-L1 and 3T3F442A cells are the most commonly used adipocyte

models for obesity studies (Ruiz-Ojeda et al., 2016). In this study we used the 3T3-L1 cell line, a well-established pre-adipocyte cell line derived from murine swiss 3T3 mouse embryos (Green & Meuth, 1974). 3T3-L1 cells exhibit a fibroblast-like morphology and upon stimulation with adipocyte differentiation inducers (insulin, dexamethasone and 3-isobutyl 1-methylxanthine (IBMX)), are converted to mature adipocytes (Green & Meuth, 1974; Green & Kehinde, 1975). Although mature 3T3-L1 adipocytes exhibit the physiological characteristics and molecular mechanisms of adipocytes in vivo, there have been concerns about how closely they mimic the pathophysiology of obesity and its associated morbidities (oxidative stress, mitochondrial dysfunction, inflammation, etc.) in vivo. Therefore, it is imperative to develop a 3T3-L1 adipocyte model that closely mimics the pathophysiology of obesity in order to allow the screening and implementation of safe and effective therapeutics to curb the burgeoning obesity epidemic.

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24

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25

3. Materials and methods

3.1 Study design

The experimental overview is illustrated in Figure 3.1. Firstly, to develop a model of increased lipid accumulation, basal lipolysis, oxidative stress and inflammation, 3T3-L1 pre-adipocytes were differentiated in 5.5 mM, 25 mM or 33 mM glucose concentrations for 7 or 14 days. Lipid accumulation, basal lipolysis, oxidative stress and inflammation were assessed using Oil Red O (ORO) staining, glycerol release, cytokine secretion (tumour necrosis factor-alpha (TNFα), MCP1 and interleukin-6 (IL6)) and quantifying ROS using the 2',7'-dichlorfluorescein-diacetate (DCFH-DA) fluorescent dye, respectively. In addition, mitochondrial dehydrogenase activity was assessed using the 3- [4, 5-Dimethylthiazol-2-yl]-2, 5 diphenyltetrazolium bromide (MTT) assay, and the expression of genes associated with lipid accumulation, basal lipolysis, inflammation and oxidative stress were measured using quantitative real-time polymerase chain reaction (qRT-PCR).

Secondly, the anti-obesity effects of two plant extracts and their major polyphenols were assessed by investigating their ameliorative properties against lipid accumulation, basal lipolysis, oxidative stress and inflammation in the optimised experimental model developed previously. Briefly, 3T3-L1 adipocytes were treated with a standardised green rooibos extract of Aspalathus linearis (GRT), a crude polyphenol enriched fraction of Cyclopia intermedia (CPEF) and their major compound Aspalathin and Mangiferin, respectively. The treatment was divided into acute and chronic treatment, whereby mature adipocytes were treated for 24 hrs or 7 days, respectively.

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26 Figure 3.1 Experimental overview

3.2 Materials

The murine 3T3-L1 pre-adipocyte cell line (ATCC® CL-173™) was purchased from the American Type Culture Collection (ATCC), Manassas, VA, USA. GRT, a standardised, commercial extract of Aspalathus linearis containing 12.8% of Aspalathin was obtained from Afriplex (Pty) Ltd., Paarl, Western Cape, SA. The CPEF was supplied by Babalwa Jack (Jack et al., 2017). Aspalathin (ca. 98% purity) was purchased from High Force Research Ltd., Durham, UK. Mangiferin (≥ 98% purity) was purchased from Sigma-Aldrich, St Louis, MO, USA and Isoproterenol was purchased from Merck (Sigma-Aldrich, St Louis, MO, USA). All other chemicals were purchased from Sigma-Aldrich, St Louis, MO, USA, unless otherwise stated. All materials, suppliers and product numbers are listed in the Appendix.

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27 3.3 Cell culture

Cell culture was conducted adhering to aseptic technique (Appendix). The 3T3-L1 pre-adipocytes were cultured and maintained in Dulbecco’s modified eagle media (DMEM, Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 25 mM glucose and 10% fetal bovine serum (FBS, Gibco, Thermo Fisher Scientific, Waltham, MA, USA) (standard conditions for 3T3-L1 culture recommended by the ATCC) (ATCC, 2013), or in 5.5 mM, 25 mM, or 33 mM glucose and 10% FBS for experimental conditions. Cell culture procedures were conducted in a biohazard safety cabinet, class II (Airvolution lab, Johannesburg, Gauteng, SA) and cells were maintained in an incubator (Galaxy R CO2 incubator, RS Biotech, West Lothian, UK) at 37°C in humidified air

with 5% carbon dioxide (CO2) (Air Products, Bellville, Western Cape, SA).

3.3.1 Thawing and culturing of 3T3-L1 cells

A cryogenic vial containing 3T3-L1 cells (1×106 cells/ml) in cryopreserving medium containing DMEM, 10% FBS and 7% (v/v) dimethyl sulfoxide (DMSO) (Sigma-Aldrich, St Louis, MO, USA) was removed from liquid nitrogen storage and thawed in a 37°C water bath. Immediately after thawing, the cells were transferred into a 15 ml centrifuge tube containing 9 ml of 37°C pre-warmed, pre-adipocyte growth medium (DMEM supplemented with 10% FBS) and centrifuged at 800 × g for 5 min (Eppendorf 5810 Centrifuge, Eppendorf, Hamburg, Germany). Thereafter, the supernatant was aspirated, and the cell pellet was resuspended in 5 ml of warmed, pre-adipocyte growth medium. Following this, 1 ml of cell suspension was transferred to a 75 cm2

tissue culture flask (T75) (Nest Scientific, Rahway, NJ, USA) containing 17 ml of pre-warmed, pre-adipocyte growth medium. The cells were incubated under standard cell culture conditions (37°C in humidified air with 5% CO2) for 24 hrs, where after the pre-adipocyte growth medium

was refreshed. Cells were kept under standard cell culture conditions until they reached 70-80% confluency and subsequently sub-cultured using a split ratio of 1:5 (section 3.3.2). Passage number was kept to below 20 in order to prevent the depletion of specific cell phenotypes such as the ability of these cells to differentiate due to excessive passage and to maintain consistency between experiments.

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28 3.3.2 Sub-culture of 3T3-L1 cells

3.3.2.1 Trypsin treatment

Upon confluency, pre-adipocyte growth medium was aspirated from the T75 flask, (Nest Scientific, Rahway, NJ, USA) and cells were rinsed with pre-warmed Dulbecco’s phosphate buffered saline (DPBS) (Lonza, Walkersville, MD, USA). Thereafter, the adherent cells were dislodged by incubating with 2 ml of trypsin (Lonza, Walkersville, MD, USA) under standard cell culture conditions for 5 - 7 min. Cells were viewed under an Olympus inverted light microscope (CKX 41, Olympus; Melville, NY, USA) to confirm whether they had dislodged from the flask. Trypsinisation was stopped by adding 8 ml of pre-warmed pre-adipocyte growth medium directly to the cells, where after the cell suspension was thoroughly mixed by gently pipetting up and down at least 8 times to disaggregate cell clumps and ensure a single cell suspension. Thereafter, the cell suspension was transferred to a 15 ml centrifuge tube and centrifuged for 5 min at 800 × g (Eppendorf 5810 Centrifuge, Eppendorf, Hamburg, Germany). The supernatant was aspirated, and the cell pellet was resuspended in pre-warmed, pre-adipocyte growth medium, subsequently sub-cultured in T75 flasks and incubated under standard cell culture conditions. Cell density and viability were determined (section 3.3.2.2) prior to freezing cells for storage (section 3.3.2.3) or seeding cells in their appropriate multi-well plates for subsequent in vitro bioassays (section 3.3.2.4).

3.3.2.2 Cell counting

Following trypsinisation, the number of viable cells were quantified using a haemocytometer and trypan blue dye (Invitrogen, Carlsbad, CA, USA). Briefly, a 10 μl volume containing the cell suspension and trypan blue in a 1:1 ratio was loaded onto one of the haemocytometer chambers (Figure 3.2B) and viewed under an inverted microscope where both viable (clear/unstained) and non-viable (stained blue) cells were counted in four of the nine quadrants of the haemocytometer (Figure 3.2C). The percentage of viable cells was calculated using the formula in Figure 3.2A. If the viability of cells was 70% or above, they were used for making stocks for freezing or seeded into multi-well plates for subsequent in vitro bioassays.

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29 Figure 3.2 Cell counting

Equation for determining cell viability (A), an illustration showing the haemocytometer slide with two counting chambers (B) and the number of squares used for cell counting (indicated by grey circled numbers) to obtain number of cells/ml (C). Cells on the bottom and right lines of the quadrant were not counted.

3.3.2.3 Freezing 3T3-L1 pre-adipocytes

Subsequent to cell counting, cells were centrifuged at 800 × g for 5 min. The supernatant was carefully discarded without disturbing the pellet, and the pellet was resuspended in sterile cold freezing medium containing DMEM, 10% FBS and 7% (v/v) DMSO at a volume required to achieve 1 × 106 cells/ml. The resuspended cell suspension was aliquoted into cryotubes (1 ml) and placed on ice. Each cryovial was labelled with the cell line, passage number, cell concentration and date. Cells were stored overnight at -80°C and thereafter transferred to liquid nitrogen for long term storage.

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30 3.3.2.4 Seeding of 3T3-L1 pre-adipocytes

After quantifying the number of viable cells, 3T3-L1 pre-adipocytes were seeded at relative seeding densities into their respective multi-well plates (Table 3.1). The seeding cell concentration for 3T3-L1 pre-adipocytes is 2 × 104 cells/ml. Thereafter, the cells were incubated under standard cell culture conditions in pre-adipocyte growth medium, until they reached 100% confluency (~4 days) and the medium was refreshed every 48 hrs.

Table 3.1 Cell densities used for seeding 3T3-L1 pre-adipocytes Cell culture plates Seeding Concentration (cells/ml) Cell density (cells/well) Volume (ml) 96-well 2 × 104 4 × 103 0.2 ml 24-well 2 × 104 2 × 104 1 ml 6-well 2 × 104 _{6 × 10}4 _{3 ml} 3.3.3 3T3-L1 pre-adipocyte differentiation

Adipocyte differentiation was conducted in accordance with the ATCC protocol (ATCC, 2013), with slight modifications in order to develop an in vitro 3T3-L1 adipocyte model with increased lipid accumulation, basal, oxidative stress and inflammation. Briefly, fully confluent 3T3-L1 pre-adipocytes (100% confluent at day 4 post seeding) were induced to differentiate into pre-adipocytes by incubating the cells (day 0) in adipocyte differentiation medium (ADM) consisting of DMEM, at various concentrations of glucose (5.5 mM, 25 mM and 33 mM) each supplemented with 10% FBS, 0.5 mM IBMX (Sigma-Aldrich, St Louis, MO, USA), 1 µg/ml insulin (Sigma-Aldrich, St Louis, MO, USA) and 1 μM dexamethasone (Sigma-Aldrich, St Louis, MO, USA ) under standard cell culture conditions until day 3 of differentiation (Figure 3.3). After 72 hrs (day 3), the ADM was changed to adipocyte maintenance medium (AMM) consisting of DMEM, at various concentrations of glucose (5.5 mM, 25 mM and 33 mM) each supplemented with 10% FBS and 1 μg/ml of insulin for a further two days (Figure 3.3). On day 5, adipocytes were incubated in DMEM

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31 (5.5 mM, 25 mM or 33 mM glucose concentrations) supplemented with 10% FBS under standard cell culture conditions until day 7 or day 14 of differentiation, with medium refreshed every 48 hrs (Figure 3.3). The cells and the cell culture media were collected at days 0, 7 and 14 of differentiation, and used for bioassays and gene expression analysis as shown in Figure 3.3.

3.3.4 Cell culture media collection

After the 3T3-L1 cells were differentiated or treated as described in sections 3.3.3 and 3.3.5, cell culture media were collected into 15 ml centrifuge tubes (Nest Scientific, Rahway, NJ, USA). The collected media were centrifuged (SL 16R Thermo Fisher Scientific, Waltham, MA, USA) at 3 500 × g for 15 min at 4°C to remove cell debris. Thereafter, the media were aliquoted and stored at - 80°C, for subsequent measurement of basal lipolysis (section 3.6) and inflammation (section 3.8).